self-replenishing vascularized fouling-release surfacesself-replenishing vascularized...

Self-Replenishing Vascularized Fouling-Release SurfacesCaitlin Howell,*,†,‡ Thy L. Vu,†,‡ Jennifer J. Lin,†,‡ Stefan Kolle,†,‡ Nidhi Juthani,†,‡ Emily Watson,†,‡

James C. Weaver,† Jack Alvarenga,† and Joanna Aizenberg*,†,‡,§

†Wyss Institute for Biologically Inspired Engineering, 60 Oxford Street, Cambridge, Massachusetts 02138, United States‡School of Engineering and Applied Sciences, Harvard University, 29 Oxford Street, Cambridge, Massachusetts 02138, United States§Department of Chemistry and Chemical Biology and Kavli Institute for Bionano Science and Technology, Harvard University, 12Oxford Street, Cambridge, Massachusetts 02138, United States

ABSTRACT: Inspired by the long-term effectiveness of livingantifouling materials, we have developed a method for the self-replenishment of synthetic biofouling-release surfaces. Thesesurfaces are created by either molding or directly embedding3D vascular systems into polydimethylsiloxane (PDMS) andfilling them with a silicone oil to generate a nontoxic oil-infused material. When replenished with silicone oil from anoutside source, these materials are capable of self-lubricationand continuous renewal of the interfacial fouling-release layer.Under accelerated lubricant loss conditions, fully infused vascularized samples retained significantly more lubricant thanequivalent nonvascularized controls. Tests of lubricant-infused PDMS in static cultures of the infectious bacteria Staphylococcusaureus and Escherichia coli as well as the green microalgae Botryococcus braunii, Chlamydomonas reinhardtii, Dunaliella salina, andNannochloropsis oculata showed a significant reduction in biofilm adhesion compared to PDMS and glass controls containing nolubricant. Further experiments on vascularized versus nonvascularized samples that had been subjected to accelerated lubricantevaporation conditions for up to 48 h showed significantly less biofilm adherence on the vascularized surfaces. These resultsdemonstrate the ability of an embedded lubricant-filled vascular network to improve the longevity of fouling-release surfaces.

KEYWORDS: fouling-release surfaces, antifouling, self-replenishing, PDMS, bacteria, algae

■ INTRODUCTION

Biofouling is a dynamic, complex process. In nature, foulingorganisms have developed extremely effective strategies toattach themselves to a wide variety of surfaces under diverseand frequently changing environmental conditions. For humanbeings, such effectiveness and adaptability of fouling organismshas proven to have significant negative economic andenvironmental consequences across a wide variety of areas.Biofilm formation on medical devices is responsible for half ofall healthcare-associated infections,1 costing U.S. hospitalsbetween $28 and $45 billion annually2 and unquantifiableloss in quality of life for many patients. In ships, biofouling canresult in power penalties of anywhere between 10 and 86%,3

and has been estimated to cost between $180−260 million peryear for the U.S. Navy fleet alone.4 Biofouling also impacts foodpackaging and storage, industrial-scale growth and release ofalgae for biofuels and other products, and water purificationsystems, to name just a few.5−8

Many strategies have been developed to reduce thedetrimental impacts of biofouling. Current approachesgenerally rely on either chemical modifications, structuralmodifications, or some combination of the two. Chemicalapproaches include hydrophilic coatings such as polyethyleneglycol (PEG)9,10 that prevent nonspecific adhesion of bacteriaand proteins, as well as substrates that incorporate or releasematerials toxic to organisms, such as quaternary ammonium

salts and silver nanoparticles in medical environments,11 ortributyltin (TBT) and tributyltin oxide (TBTO) in marinesettings.12 Physical approaches typically use micro- or nanoscalesurfaces to reduce organism contact area and adhesionstrength.13−16 One such method, slippery liquid-infused poroussurfaces (SLIPS), makes use of a surface-immobilizedomniphobic lubricant layer to prevent the adherence of foulingorganisms as well as other materials.17−21 Yet although many ofthese treatments show promise on short time scales, nearly allwill begin to fail over extended periods. Chemical coatings aresubject to degradation, depletion, or desorption,22 and physicalmodifications can be structurally damaged thus creating sitesfor adhesion. Furthermore, in many cases, fouling organismsthemselves adapt by generating a conditioning layer of proteinsor polysaccharides to mask and condition the underlyingsurface.22,23

Fouling-release coatings are an alternative approach that hasthe potential to be used over longer periods of time. Currentlyused in marine environments as nontoxic alternatives tobiocidal paints,12,24,25 these coatings are based on the conceptthat physical removal of fouling agents appears to be easiestfrom materials with low elastic moduli and surface free energies

Received: May 21, 2014Accepted: July 9, 2014Published: July 9, 2014

Research Article

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between 20 and 30 mJ/m2,24,26 such as silicones. Foulingorganisms such as barnacles, algae spores, and bacteria cantherefore be removed from fouling-release surfaces with theapplication of relatively low hydrodynamic forces.27 It has alsobeen observed that the addition of some oils to siliconepolymer coatings prior to curing can further enhance thefouling-release capabilities of these surfaces28 and decreasefouling levels even after 2 years in marine field tests.29 Yet eventhese materials will eventually become fouled.In nature, antifouling strategies adopted by various plants and

animals not only successfully prevent the attachment andproliferation of fouling organisms, but they continue to do soover weeks, months, and even years, stopping only when theorganism is dead. Various adaptations, such as complexnanoscale surface topographies on the skin of sharks andpilot whales, coatings of the fish scales with the layer of mucus,or the production of natural biocidal chemicals on the fronds ofmacroalgal species, e.g. Delisea pulchra, allow these organismsto remain clean over extended periods of time.30 Yet althoughmuch effort has been put into duplicating these effects bymimicking surface structure or chemical composition of thesenatural systems, no synthetic antifouling material has proven tobe as effective as shark and fish skin or algae fronds when itcomes to long-term functionality.30 The reason for this is atleast partly due to the fact that surfaces on living animals andplants are dynamic, just as the organisms that try to foul them.Living surfaces can be constantly sloughed off and replenished

to remove or discourage biofoulers, as opposed to staticsurfaces that can almost always be overcome given enoughtime. Some previous work has been done to try to mimicdynamic living surfaces by developing skin-like coatings thatdissolve in marine environments to release fouling agents.31

However, many technical challenges related to the durabilityand scale-up of these types of materials remain.Here, we describe the creation and initial testing of fouling-

release surfaces made self-replenishing through the incorpo-ration of a vascular system as an internal reservoir. We startfrom a common silicone-based fouling-release material(polydimethylsiloxane [PDMS]), and infuse it with siliconeoil to improve its release properties. Finally, we fill theunderlying vascular network with more silicone oil lubricant,which can diffuse through the solid PDMS to continuallyreplenish the interfacial lubricating layer. We describe twomethods for fabricating these vascular networks and demon-strate capacity of the system for lubricant retention andreplenishment in accelerated tests. We further show thatlubricant-infused silicone polymers display excellent fouling-release properties against bacterial monocultures and algaemixed cultures, and that these properties can be retained overtime through the incorporation of lubricant-containing vascularnetworks. This work represents the next step toward thedevelopment of true bioinspired “living” nontoxic antifoulingcoatings.

Figure 1. (A) Schematic of the process to make either vascularized or nonvascularized silicone-oil-infused PDMS. For simple infused PDMS (upperrow), cured PDMS is placed in a bath of silicone oil, which diffuses into the PDMS solid. For vascularized PDMS, the vascular pattern is createdbefore curing. The sample is then infused externally with silicone oil in the same manner as the nonvascularized PDMS, or internally through fillingthe vascular network, or both. (B) Methods of creating vascular networks within PDMS: (1) An encased network is created using a 3D mold. (a−e)The schematic of the procedure. The target pattern is molded (a) to create the pattern in PDMS. (b) The mold is removed from the cured PDMS(c) and the pattern is covered with a second sheet of PDMS (d), which is chemically bonded to the pattern using O2 plasma (e). (i) An image of a3D leaf vasculature network after encasing. (2) An embedded network is created following the procedures developed by Lewis et al.32 (a) A patternof 20% w/w Pluronic F127 gel at 25 °C is embedded in uncured PDMS. (b) The PDMS is allowed to cure, cooled to 4 °C, and the liquid Pluronic isevacuated. (c) The channel is refilled with silicone oil. An image of fluorescently dyed silicone oil in (i) a sinusoidal channel, (ii) a leaf-shapenetwork, and (iii) a linear network.

ACS Applied Materials & Interfaces Research Article

dx.doi.org/10.1021/am503150y | ACS Appl. Mater. Interfaces 2014, 6, 13299−1330713300

■ EXPERIMENTAL SECTIONFabrication of PDMS Vascular Systems (Figure 1). Sylgard 184

silicone elastomer (Dow Corning Corporation, Midland, MI) was usedto fabricate all samples used in the experiments described. The baseand curing agent were combined in a 10:1 ratio and mixed in a Thinkyplanetary centrifugal mixer (Thinky Corporation, Tokyo, Japan) at2000 rpm for 1 min, then again at 2200 rpm for 1 min.Encased Vasculature. Vascularized masters were fabricated by 3D

printing using a Stratasys Connex500 3D printer (Stratasys Ltd., EdenPrairie, MN) with VeroBlack polyjet resin. 2D images of leaf vascularsystems were employed as the original input files. Prior to molding, themasters were kept at 100 °C for 24 h. Uncured PDMS was thenpoured onto the 3D printed master, degassed under vacuum, andallowed to cure at 70 °C for at least 2 h. Once curing was complete,the mold was carefully separated from the master. Finally, thevasculature was encased by chemically bonding a second sheet ofPDMS over the mold by applying oxygen plasma (30 s) followed byheat treatment (2 h at 70 °C) to both surfaces (Figure 1B(1)).Embedded Vasculature. Channels were embedded into PDMS

according to procedures developed by Lewis et al.32 Briefly, a 20% (w/v) solution of Pluronic F127 (Sigma-Aldrich, St. Louis, MO) dissolvedin water was heated until it became a gel (normally around 70 °C) andmanually embedded into uncured, degassed PDMS in desired channelpatterns. The PDMS was then allowed to cure completely at 70 °C.Finally, the PDMS samples were cooled to 4 °C to cause the PluronicF127 solution within the channels to become liquid. The liquid wasthen evacuated using an in-house vacuum system, leaving empty,embedded channels (Figure 1B(2)).Lubricant Infusion. The channels in the vascularized PDMS were

filled with Momentive Element 14* 10A silicone oil (Momentive,Albany, NY). These samples, as well as the unvascularized PDMSsamples for controls and initial biological tests, were completelyimmersed in a silicone oil bath for 2−20 days (depending on initialthickness) until the mass of the samples no longer increased and fullinfusion of the lubricant into the PDMS molecular structure hadoccurred.Lubricant Replenishment Tests. Accelerated testing of lubricant

replenishment in cylindrical vascularized samples compared tononvascularized controls was performed by placing fully infusedsamples in an oven at 70 °C. Cylindrical embedded vascularizedsamples and equivalent controls were used due to their smaller sizeand therefore faster rate of infusion. In the vascularized samples, thefree silicone oil took up approximately 17% of the entire samplevolume. Samples were gently wiped prior to the experiment to removeany excess lubricant from the surface. The masses of the samples weretaken daily using a Mettler Toledo Xs205 DualRange analyticalbalance to measure lubricant loss via evaporation. Before weights weretaken, the channels of the vascularized samples were refilled using asmall-gauge syringe inserted through the encasing PDMS into thechannel in one corner of the sample. The air was allowed to escape viaa second small puncture in the opposite corner. This was done inorder to mimic a system with a continuous lubricant supply.Bacterial Biofouling Tests. Liquid stock solutions of Staph-

ylococcus aureus strain SC-01 in tryptic soy broth (TSB) medium andEscherichia coli strain J96 in M63 medium with densities ofapproximately 1 × 108 cells/mL were prepared. S. aureus and E. coliwere chosen because of their role as an infection-causing agents inmany healthcare settings.1 The stock solutions were diluted 1:100 (v/v) in either M63 or TSB biofilm medium (TSB with 1.5% (w/v)NaCl) in Petri dishes containing samples of glass, noninfused PDMS,and lubricant-infused PDMS (nonvascularized). Cultures were thenincubated at 37 °C for 48 h.Following incubation, the substrates were gently removed from the

biofilm medium. Samples for biofilm coverage analysis wereimmediately stained by incubating in a 0.1% (w/v) solution of crystalviolet in water for 10 min, gently rinsed in deionized, distilled water toremove excess stain, then dried and photographed. Samples forcolony-forming unit (CFU) counts were sonicated for 5 min in 1xphosphate buffered saline (PBS) to remove the bacteria from the

surfaces, then diluted and plated out on TSB agar. For both biofilmcoverage and CFU counts, at least five replicates per treatment wereprepared.

For bacterial biofouling tests on vascularized PDMS versus controls,samples were kept under static conditions at 70 °C for 1, 2, 4, 8, 24,and 48 h to cause lubricant loss through evaporation. Bothunlubricated samples and fully lubricated samples that had not beenexposed to evaporation were included as controls. The channels of thevascularized samples were kept continuously filled in the same mannerused for the lubricant replenishment tests. Samples were thenimmersed in bacterial medium for 48 h, removed, and stained withcrystal violet as described above.

Algae Biofouling Tests. Stock solutions of nonaxenic Botryococcusbraunii (UTEX number 2441)33 and Chlamydomonas reinhardtii(UTEX number 89) from the University of Texas Culture Collectionwere grown in Basal Medium and Bristol Medium, respectively.Dunaliella salina cultures were provided by the Hastings Lab atHarvard University and maintained in Guillard f/2 marine waterenrichment solution (50x) (G0154, Sigma-Aldrich, St. Louis, MO).Nannochloropsis oculata cultures were provided by Solix Biosystemsand grown in a modified f/2 medium with vitamins omitted.

Once a density of approximately 1 × 107 algal cells per mL wasreached, these cultures were diluted 1:5 in Petri dishes containing freshmedium. These dishes contained noninfused and lubricant-infusedPDMS (nonvascularized) test surfaces created by spin-coating PDMSonto flat glass substrates, as well as glass controls. Cultures wereincubated under a set of Sun Blaze T5HO fluorescent light fixtures(Sunlight Supply, Inc., Vancouver, WA). The lamps were on timers toprovide 16 h of light followed by 8 h of darkness. Surface temperatureswere monitored throughout the experiment and did not exceed 25 °C.There were at least five replicates per treatment.

After 12 days, the substrates were gently lifted at one end to attach aclip connected to a dip-coater via a nylon cord. The substrates werethen lifted out of the culture medium at a controlled rate of 0.5 mm/s.Once the substrate was completely clear of the medium, it wasphotographed for image analysis. Immediately after photographing, thesamples were processed for chlorophyll a as a proxy for estimatedbiofilm mass. Each sample was placed into a 600 mL beaker filled with300 mL of distilled, deionized water (DIW) and sonicated for 2 min.The resulting solution biomass was then vacuum-filtered using aceramic funnel with Whatman 1 filter papers. Each filter paper wasimmersed in 5 mL DMSO and was heated overnight at 70 °C. A 200μL volume of this solution was then measured for absorbance at 665and 750 nm. The resulting chlorophyll a values were then determinedusing the following equation for each sample34

α= − −C D D R R v l( )[ /( 1)]( / )(10 / )a b a3

c

where Ca is the concentration of chlorophyll a in mg/mL solvent, Dband Da are the optical densities before and after acid addition,respectively, αc is the specific absorption coefficient for chlorophyll a, vthe volume of solvent used to extract the sample, and l the path lengthof the spectrophotometer in cm. R is defined as Db/Da for purechlorophyll a. The values obtained at 750 nm were subtracted fromthose at 665 nm to account for sample turbidity.34

Toxicity Tests. Experiments to determine toxicity were conductedon B. braunii cultures as these were considered likely to be moresensitive to adverse effects from silicone oil (suffocation, lightinterference) than bacteria due to their need to photosynthesize.27 A500 mL volume of Basal Medium containing a 1:5 ratio of B. brauniistock was placed into 1 L Erlenmeyer flasks either with or without 10%(v/v) of additional silicone oil. The cultures were then bubbled withair to ensure continuous mixing. There were five replicates pertreatment.

Every 3 days, approximately 1 mL of the culture medium wasremoved for analysis. Of this, a 500 μL aliquot was incubated with 10μM Sytox Green Nucleic Acid Stain (Life Technologies, Carlsbad,CA) for 10 min, then visualized using a Zeiss microscope with anexcitation wavelength at 470 nm.35 Using this procedure, all dead cellswere stained green. Live algae produced red autofluorescence due to



their chlorophyll a content. Five images were taken per sample. Cellswere counted using MATLAB (Mathworks, MA, USA) prior tostatistical analysis. A 200 μL aliquot of the remaining material was usedto measure optical density at 683 nm using a SpectraMax 384 platereader (Molecular Devices, LLC, Sunnyvale, CA).Biofilm Coverage Analysis. After removal from the liquid

medium (and crystal violet staining in the case of the bacterial foulingtests), all test surfaces were photographed with a Canon EOS RebelT4i camera (Melville, NY). Images were analyzed by thresholding suchthat biofilm-covered areas became white and the clear areas becameblack. This allowed for the calculation of the area that was thresholdedand hence the area of the biofilm, which was divided by the area of thesample to determine the percent coverage. Image analysis wasperformed either using ImageJ or MATLAB (Mathworks, MA, USA).Statistics. For image analysis, the percent area values between 0

and 100% were first arcsine transformed to change the distributionfrom binomial to nearly normal.36 Two-tailed t tests or univariateAnalyses of Variance (ANOVAs) with Tukey posthoc test were thenperformed to determine statistical significance between the control andlubricant-infused treatments. Live cell counts in the toxicity tests wereanalyzed using a repeated-measures two-way ANOVA. All statisticalanalyses were performed using IBM SPSS Statistics version 20 (IBM,Armonk, NY). A value of P < 0.05 was considered significant.

■ RESULTS AND DISCUSSION

Fabrication of PDMS Vascular Systems. Samples ofeither vascularized or nonvascularized lubricant-infused PDMSwere prepared as schematically depicted in Figure 1A. Vascularsystems were fabricated either as encased channels orembedded channels as described in the materials and methodssection. Figure 1B(1) shows the process of fabricating anencased vasculature system based on the vein structure of a leaf.The desired 2D pattern was converted into a black and whitevector drawing, imported into SolidWorks (Dassault Systems,Waltham, MA), then 3D printed to make a master. The masterwas used to mold the vascular pattern into PDMS. The finalmold was then covered in a second sheet of PDMS to enclosethe vascular channels. With this fabrication method it waspossible to create complex channel systems over a potentiallywide area (depending on the size of the 3D printed master).However, it was noted that great care needed to be taken whenattaching the covering PDMS sheet to the final mold. Anyincomplete chemical bonding between the two PDMS pieces,because of such factors as overexposure to plasma or presenceof surface contamination, was found to result in delaminationupon lubricant infusion.Embedding the vascular system directly into uncured PDMS

was explored as a way to avoid delamination, as well as a way tomake multiple smaller samples in parallel for further character-ization. Lewis et al.32 developed the method of using PluronicF127 for embedding 3D-printed channel systems intopolymers, such as PDMS, for culturing cells. This compound,when mixed with water at a 20% w/v ratio, produces a solutionwhich is a gel above 25 °C, but a liquid at around 4 °C. Byexploiting this property it was possible to manually drawchannels into PDMS using the gel-phase Pluronic at 25 °C,cool it to 4 °C and remove the then liquid-phase Pluronic usingvacuum, followed by refilling the channels with silicone oillubricant. Using this procedure, a variety of simple vascularsystems could be created (Figure 1B(2)).Lubricant Replenishment Tests. Experiments were

conducted on embedded vascular systems and nonvascularizedcontrols to compare their ability to secrete the lubricant fromthe channels to the surface upon interfacial lubricant loss.Evaporation at 70 °C was chosen as the test method of

lubricant removal from the surfaces as opposed to wiping ordepletion via water flow for two reasons: first, it provided amore even removal than physical wiping without damaging thesurfaces; and second, it removed lubricant from the surfacesmore quickly than the low-flow conditions, to which thesematerials would be exposed in medical devices or stationarymarine applications, thus offering a more suitable acceleratedtest. Evaporation at elevated temperatures would also beexpected to be the primary route of lubricant loss if thesematerials were exposed to alternating marine/open airconditions (such as coatings for dock pillars exposed to tides,for example), as previous experiments on radiolabeled oil lossfrom PDMS samples showed less than 1% loss over 1 yearunder simulated marine conditions when not exposed toevaporation.29

The channels containing free oil took up approximately 17%of the total volume of the vascularized samples, and samplessaturated with the lubricant oil had masses approximately 30%greater than unsaturated samples. The results of acceleratedevaporation tests on these initially saturated samples gatheredover 11 days are shown in Figure 2. The vascularized PDMS

samples lost 4.6% (±2.5%) of their lubricant volume (withcontinual lubricant replenishment), while the nonvascularizedcontrol lost 13.5% (±1.2%). There was a clear significantdifference in the amount of lubricant lost between the two (P <0.01). After 24 days, this difference had widened to 6.5%(±3.7%) lubricant loss in the vascularized samples compared to20.8% (±1.2%) loss in the controls.These results show that the presence of a vascular system

slows lubricant loss via evaporation in PDMS samples,providing on-demand replenishment of the lubricant in thesystem that can counteract lubricant losses from the surface.

Biofouling Tests. Previous studies have shown that theincorporation of free silicone oil into silicone polymer coatingscan help increase their fouling resistance in marine settings.28,29

However, these samples were generally created as one-potcoatings already containing free oil introduced during polymer-ization rather than as oil added to the polymer after curing bypostfabrication swelling or as oil diffusing to the interface frombelow. To ensure that the samples saturated with oilpostfabrication would also function efficiently as fouling-release

Figure 2. Mass of saturated lubricant-infused PDMS samplescontaining an embedded vascular network (squares) versus a samplewith no vascular network (diamonds) kept at 70 °C for 11 days. Thedata were normalized to the dry mass of the samples and error barsrepresent standard deviation. The control lost mass because oflubricant evaporation more rapidly than the channel system in thisaccelerated test.



surfaces against model organisms, we tested fully infusedPDMS substrates against static cultures of the infectiousbacterium S. aureus1 as well as against the model microalga B.braunii.33,37 These test were performed first on nonvascularizedsamples, as lack of evaporation or other lubricant-removingforces made the inclusion of vasculature unnecessary.The results of the static tests against S. aureus are shown in

Figure 3. Both glass and noninfused PDMS were used aspositive controls to demonstrate the effects of biofilmformation on hydrophilic and hydrophobic surfaces, respec-tively. Panels A and B in Figure 3 show that both of thesesurface types fail to resist biofilm attachment, resulting inbiofilm coverages between 73.9% and 64.8% (±7.5%) (Figure3C), and CFU counts of 6.5 × 108 and 10.7 × 108 cells/cm2

(Figure 3D) for glass and untreated PDMS, respectively. Thelubricant-infused PDMS samples, in contrast, showed no visibleadherent biofilms upon being removed from the culturemedium, resulting in surface coverage values of 0.1%(±0.1%) after crystal violet staining. CFU counts for thesesamples yielded values of 4.8 × 105 cells/cm2, 3 orders ofmagnitude lower than the control samples. These resultsdemonstrated that lubricant-infused PDMS can be significantlymore effective as fouling-release surfaces against biofilms of S.aureus compared to even low-adhesion surfaces of PDMS orglass.Biofouling tests on substrates in static cultures of the

microalgae B. braunii showed similar results. Upon removal ofthe substrates from the culture medium, the biofilm peeled

Figure 3. (A) Removal of glass, PDMS, and lubricant-infused PDMS from a solution of S. aureus after 48 h of growth. The black arrow indicates thelocation of the infused PDMS sample. Note that the controls become cloudy due to the biofilm overgrowth, while the sample that was infused withthe oil postfabrication is completely clear and transparent. (B) Representative samples after staining for adherent biofilm with crystal violet. A blackline is added to outline the infused PDMS sample due to its near-complete transparency. (C) Image analysis results (% biofilm coverage) of thesurfaces. (D) CFU counts showed a decrease of at least 3 orders of magnitude in the lubricant-infused PDMS samples compared to the controlPDMS and glass samples.

Figure 4. (A) Glass slides half-coated with lubricant-infused PDMS were incubated in culture vessels containing B. braunii. After 12 days, the slideswere slowly removed from the vessel using a dip coater attached to the slide. At t = 0 s and t = 5 s, the clear attachment of the algae biofilm to theuntreated glass can be seen. However, at t = 15 s the boundary between the treated and untreated class is crossed, and at t = 16 s the biofilm beginsto peel away from the surface. The arrow at t = 35 s indicates the intact biofilm floating on the surface of the culture medium. (B) Percent biofilmsurface coverage of the glass substrates, untreated PDMS substrates, and lubricant-infused PDMS substrates after removal. (C) Amount ofchlorophyll a remaining on the surfaces after removal.



nearly completely off of the lubricant-infused samples, while itremained firmly attached on the control samples. This effectwas most striking on glass substrates that had been half-coatedin lubricant-infused PDMS, as shown in Figure 4A. As thesesamples were removed from the culture solution, the biofilmsbegan to peel away from the surface as soon as the lubricant-infused coating was reached. In fact, it is interesting to note thatthe biofilm was so poorly adhered to the lubricant-infusedsurface compared to the glass surface that the film was almostperfectly ripped in half at the glass/lubricant-infused PDMSboundary. This effect is most difficult to obtain in the directionof removal shown in Figure 4A (first glass, then lubricant-infused PDMS), as the biofilm initially covered the entiresurface and would have therefore offered some stability acrossthe boundary. After removal of the surfaces, the intact algaebiofilm could be observed floating on the surface of the culturemedium (as indicated by a white arrow).Some adherent biofilm did remain on the edges of the

lubricant-infused half of the sample. This was due to edgeeffects of the spin-coated PDMS providing areas of incompletesurface coverage. Such remaining material would be minimizedin larger coated samples or materials made exclusively oflubricant-infused polymer. Analysis of percent biofilm coveragefor completely coated surfaces yielded values of 99.8% and94.5% (±0.7%) for the glass and PDMS substrates, respectively,while for the lubricant-infused PDMS surfaces this value was aslow as 6.6% (±6.4%) even with the contribution from the edgeeffect (Figure 4B). Quantification of the amount of chlorophylla on these surfaces gave 269.5 and 243.6 μg/mL (±86.2 μg/mL) for glass and PDMS, respectively, compared to 13.4 μg/mL (±12.2 μg/mL) for the lubricant-infused PDMS (Figure4C). These data confirmed that lubricant-infused PDMS wasalso effective at releasing mixed microalgae/bacterial biofilms.Furthermore, taken together with the S. aureus data, the resultsshowed that for these types of lubricant-infused substrates allthat was needed to remove the adherent biofilm was the smallamount of force applied by passing the samples through theair/water interface.Similar experiments were performed using the three

commercially relevant species C. reinhardtii, D. salina, and N.oculata, along with their naturally associated bacteria, toevaluate the effectiveness of PDMS that had been infusedwith oil postfabrication when exposed to different algae species.B. braunii was selected for initial studies as it forms strongbiofilms under static conditions and is also well-known for itscapacity to produce a very high lipid content within its cellwalls, an important factor in the cost-effective production ofbiofuels, while C. reinhardtii, D. salina, and N. oculata wereselected for their industrial relevance.38 Glass samples half-covered with lubricant-infused PDMS were placed in shallowPetri dishes containing either static (C. reinhardtii and D.salina) or shaking (N. oculata) algae cultures. The C. reinhardtiiand D. salina cultures remained green throughout theexperiment, while the N. oculata cultures became stressed andyellowed, as was expected due to the short light path.39,40 After12 days of growth, the samples were lifted out of the mediumsuch that the glass control and the lubricant-infused PDMSportions of the sample were side-by-side. The same relativelyclean separation of the biofilms along the glass/infused PDMSboundary observed for B. braunii (Figure 4) also occurred inthese samples (Figure 5).All three species tested showed significantly less surface

coverage of biofilm on lubricant-infused surfaces compared to

untreated glass control surfaces (Figure 5A). In culturescontaining C. reinhardtii, the surface coverage was 99% ± 1%for glass control surfaces compared to 16% ± 21% for lubricant-infused PDMS. The high level of variation in these samples wasdue to the presence of edge effects (see Figure 5B), similar towhat was observed in the B. braunii samples (Figure 4A).Cultures with D. salina showed biofilm surface coverage of 70 ±13% for glass controls and 4% ± 2% for lubricant-infusedsurfaces. The substrates in N. oculata cultures were stained withcrystal violet to improve contrast upon removal. The lack ofcontrast initially in these biofilms was suspected to be due tothe low chlorophyll a and high carotenoid content, as well asthe likely higher levels of bacteria that can arise in stressed algaecultures.39,40 After staining and rinsing, these surfaces showedcoverages of 74% ± 9% for glass controls and 2% ± 2% forlubricant-infused PDMS.

Fouling Tests on Vascularized Samples. To test theability of an incorporated vasculature to maintain these fouling-release effects in extreme lubricant-removal conditions, weexposed vascularized and nonvascularized substrates, which hadbeen kept at 70 °C for either 1, 2, 4, 8, 24, or 48 h to acceleratelubricant loss via evaporation, to bacterial culture. Controlsamples were found to lose up to ∼5% of their total mass (upto ∼11% of infused lubricant mass) after 48 h. The internalreservoirs of the vascularized samples were refilled at each timepoint using a syringe as described in the Experimental Section.Following evaporation, samples were exposed to E. coli culturesfor 48 h. E. coli was chosen for these experiments instead of S.aureus as the former was observed to produce more stronglyadherent biofilms on surfaces with minimal amounts oflubricant. This is not surprising, as previous work by Levkinet al. has shown that even different strains of the samebacterium can result in different biofilm attachment abilities onlubricant-infused surfaces.20

Figure 6 shows representative samples and their quantifica-tion for percent biofilm coverage after removal and stainingwith crystal violet. Unlubricated control samples (Dry) versusunevaporated samples (0h) (Figure 5A) showed results similarto what was observed for S. aureus cultures (Figure 3), e.g., a

Figure 5. (A) Surface coverage of biofilm remaining on lubricant-infused PDMS versus glass control surfaces after incubation in andremoval from the liquid medium containing C. reinhardtii, D. salina, orN. oculata. For all three species, there was a significant reduction in theamount of biofilm remaining on the lubricant-infused surfaces (P <0.05). (B) Images of the substrates used in the analysis. The biofilmsof N. oculata and associated bacteria were dyed with crystal violet inorder to improve contrast.



significant decrease in biofilm attachment on the lubricant-infused samples (P < 0.01).As the amount of time during which the samples were

exposed to lubricant loss via evaporation increased, the amountof biofilm that was able to adhere to the control samples alsoincreased, reaching a level equal to that of the unlubricatedcontrols after 48 h (Figure 6A). The vascularized samples,however, showed no significant increase in biofilm attachmentafter exposure to the same conditions because of the transportof oil from the channels to the interface.These results show that the inclusion of a vascular system can

effectively prolong the fouling-release effects of lubricant-infused PDMS even at harsh conditions of accelerated lubricantremoval.Lubricant Toxicity Tests. Although silicone oil has been

shown in multiple studies to be nontoxic to both macro- andmicro-organisms in the traditional sense,41,42 there has beensome concern that this low-surface-energy liquid could wraparound a single-celled organism, resulting in suffocation.27 Toverify that the lubricant-infused PDMS surfaces were not

showing superior biofilm release as a result of suffocation, twoseparate analyses were done to monitor the growth of B. brauniiexposed to excess amounts of the lubricant silicone oil. B.braunii was chosen for these tests instead of S. aureus for tworeasons: first, previous work has already reported similar studieson the toxicity of various other types of lubricants to bacteria18

with no effect found, and second, algae are more likely to showeffects of suffocation quickly because of their need to accessdissolved gases for photosynthesis.27

Figure 7 shows the growth curves of B. braunii in culturemedium containing 10% lubricant (v/v) compared to controlcultures containing no lubricant (note that 10 v % oil in culturerepresents orders of magnitude greater concentration of thelubricant than what can be released from the oil-infusedpolymer). Both the optical density (Figure 7A) and the totallive cell count (Figure 7B) were measured; the former to reflectthe changes in both the algal and naturally associated bacterialpopulations and the latter to show the changes in just B.braunii. Optical density measurements were taken at theexperimentally determined peak absorbance of 683 nm, inagreement with previously reported values.43 Both curvesshowed the expected initial lag phase followed by theexponential growth and subsequent saturation of thepopulation.37 Logistic regressions were conducted on theoptical density data to find the growth rate of the cultures.Statistical analysis showed no significant difference between thegrowth rate of B. braunii in lubricant-containing and lubricant-free control cultures across the 14-day time period. Repeated-measures two-way analysis of variance on the total live cellcounts also showed no significant difference between the cellcounts in lubricant-containing and control cultures over the 14-day period (P < 0.05).These results confirm that the apparent inability of these

biofilms to adhere to lubricant-infused PDMS surfaces is notdue to a lubricant toxicity or suffocation effect. Therefore, thebiofilm-release process is more likely caused by the replenish-ment of the lubricant on the PDMS surface and its fouling-release properties, perhaps through the continual sloughing ofthe biofilm off the surface lubricant layer.Previous work using slippery, liquid-infused porous surfaces

(SLIPS) exposed to cultures of Pseudomonas aeruginosa, E. coli,as well as S. aureus has shown that these organisms do not formadherent biofilms.18 Similarly, recent tests with marineorganisms on SLIPS have shown a decrease in settlement ofUlva linza zoospores and Balanus amphitrite larvae.21 These

Figure 6. (A) Quantification of the amount of biofilm remaining onthe surfaces compared to unlubricated controls (Dry) and samples thathad undergone no lubricant evaporation (0h). The samples withembedded vasculature show significantly less adherent biofilm at alltime points (P < 0.05). (B) Representative images of control samplesand samples with an embedded vascularized network after evaporationof the lubricant for 1, 2, 4, 8, 24, and 48 h and subsequent exposure toE. coli for 48 h (purple color is from crystal violet staining).

Figure 7. (A) Optical density measurements at 683 nm and (B) total live cell counts for B. braunii cultures containing 10% (v/v) lubricant (squares)and no lubricant (diamonds) over 14 days of growth. There were no significant differences in the rate of growth between cultures grown withlubricant and those grown without (P < 0.05).



surfaces were made from textured solids infiltrated with achemically matched fluorinated lubricant, but showed resultssimilar to what was observed in this work with silicone oil-infused PDMS. This supports the hypothesis that the presenceof a stable liquid barrier layer at the interface is at least partiallycontributing to the easy removal of the bacterial and microalgalbiofilms in these lubricant-infused samples.Although the method described here of creating lubricant-

infused PDMS with an embedded or encased vascular systemmay prolong its fouling-release functionality, it may noteffectively solve other problems frequently encountered whenusing PDMS as a fouling-release coating in marine environ-ments, such as low durability and bonding issues.44 While newapproaches of increasing the mechanical robustness of PDMScoatings are being currently considered, it may also be possibleto create vascular networks inside of other types of materials,such as commercially available amphiphilic coatings.44 As longas the the material used is capable of being infused withlubricant, such an approach could also increase longevity whilenot sacrificing durability or adhesion.

■ CONCLUSIONSWe have fabricated self-lubricating biofouling-release surfacesdesigned to mimic natural self-replenishing materials. Sampleswere created by either encasing a premolded vasculaturepattern or by directly embedding channels into uncured PDMS,followed by curing and filling the vascular systems with siliconeoil lubricant. Continual replenishment of the silicone oil in thevasculature from an outside source resulted in a surface thatcould self-lubricate upon oil depletion via simple diffusion ofthe lubricant from the embedded reservoir to the surface.Accelerated tests on lubricant removal via evaporation at 70 °Cshowed that the lubricant-infused vascularized systems lostsignificantly less lubricant over a 24-day period whencontinuously replenished with lubricant than nonvascularizedcontrols (6.5% versus 20.8%, respectively).When challenged with static cultures of S. aureus for 48 h, a

leading cause of infections in hospital settings,1 lubricant-infused PDMS samples showed biofilm coverages of only 0.1%,compared to 74% and 65% for glass and noninfused PDMSsurfaces, respectively. Colony-forming unit counts of thesesamples confirmed a decrease of at least 3 orders of magnitudein the number of bacteria remaining on the infused samplescompared to the controls. Tests on a clinically isolated strain ofE. coli showed similar results.Surfaces submerged for 12 days in static cultures of B.

braunii, a green microalga known for its potential in the algaebiofuels sector,38 showed biofilm coverages of 7% on lubricant-infused PDMS substrates as opposed to 98% and 95% for glassand noninfused PDMS, respectively. Chlorophyll a values ofthese samples likewise dropped to 13.4 μg/mL for infusedsubstrates, compared to 269.5 μg/mL for glass and 243.6 μg/mL for noninfused PDMS. Tests on the commercially usedspecies of C. reinhardtii, D. salina, and N. oculata also showedsignificantly reduced biofilm adherence after removal from thegrowth medium.Experiments on lubricant-infused samples kept under

accelerated lubricant loss conditions followed by exposure tostatic cultures of E. coli showed a significantly reduced amountof biofilm formation on the vascularized samples versusnonvascularized controls over multiple time points. After 48h of exposure to lubricant evaporation conditions, non-vascularized lubricant-infused samples showed 98% biofilm

coverage (equal to noninfused samples), while vascularizedcontrols showed only 5% coverage.Tests on the toxicity of the silicone oil used for infusion on

cultures of B. braunii revealed no difference in growth rate orlive cell numbers upon exposure to excessive amounts of thislubricant over a 14-day period. This eliminated toxicity or cellsuffocation as potential mechanisms of action for the fouling-release effects observed.These results demonstrate the ability of PDMS that is

saturated with the lubricant postfabrication to resist biofilmattachment from a range of bacterial and microalgal species.They further show that the incorporation of a vascular systemcan dynamically maintain the fouling-release properties of theselubricant-infused polymers even in the face of acceleratedlubricant loss conditions, mimicking the self-replenishingproperties of highly effective and long-lasting living antifoulingsystems employed by some animals and plants. Although alllubricant infusion in these studies was done using a syringe,these materials could easily be designed to include theincorporation of a lubricant inlet tube for continuous lubricantreplenishment from an external reservoir. Furthermore, coat-ings with incorporated vasculature could be applied dry andthen infused with lubricant through the vascular system alone,simplifying the coating process. In summary, the workpresented here offers the basis for future dynamic, nontoxicfouling-release coatings with significantly increased longevityarising from the incorporation of a vascular system thatprovides continuous diffusion of the lubricant to the interfaces.

■ AUTHOR INFORMATIONCorresponding Authors*E-mail: [email protected].*E-mail: [email protected] ContributionsThe manuscript was written through contributions of allauthors. All authors have approved the final version of themanuscript.NotesThe authors declare no competing financial interest.

■ ACKNOWLEDGMENTSThe authors thank Ron Parsons and Joel Butler at SolixBiosystems for providing the N. oculata cultures, and JohnSkutnik for the D. salina cultures. The authors also thank ShiraLehmann, Bobak Mosadegh and A. Arias Palomo for technicalassistance, Isa DuMond for culture assistance, and Ian Burgessfor editorial assistance. The leaf vasculature used for the 3Dmolds was taken by Jon Sullivan (PDPhoto.org) and obtainedfrom the public domain at Wikimedia Commons. Theinformation, data, or work presented herein was funded inpart by the Office of Naval Research under award no. N00014-11-1-0641 and by the Advanced Research Projects Agency-Energy (ARPA-E) under award no. DE-AR0000326.

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mailto:[email protected]

mailto:[email protected]

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